Download A cell wall reference profile for Miscanthus bioenergy crops

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A cell wall reference profile for Miscanthus bioenergy crops
highlights compositional and structural variations associated
with development and organ origin
Ricardo M. F. da Costa1, Sivakumar Pattathil2,3, Utku Avci2,3, Scott J. Lee4, Samuel P. Hazen4, Ana Winters1,
Michael G. Hahn2,3 and Maurice Bosch1
1
Institute of Biological, Environmental and Rural Sciences, Aberystwyth University, Plas Gogerddan, Aberystwyth, Ceredigion, SY23 3EE, UK; 2Complex Carbohydrate Research Center, The
University of Georgia, 315 Riverbend Road, Athens, GA 30602, USA; 3US Department of Energy Bioenergy Science Center, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA;
4
Biology Department, University of Massachusetts, Amherst, MA 01003, USA
Summary
Authors of correspondence:
Maurice Bosch
Tel: +44 1970 823103
Email: [email protected]
Ricardo M. F. da Costa
Tel: +44 1971 03162
Email: [email protected]
Received: 18 July 2016
Accepted: 29 September 2016
New Phytologist (2017) 213: 1710–1725
doi: 10.1111/nph.14306
Key words: bioenergy, biomass,
carbohydrate, cell wall, glycan,
lignocellulose, miscanthus, recalcitrance.
Miscanthus spp. are promising lignocellulosic energy crops, but cell wall recalcitrance to
deconstruction still hinders their widespread use as bioenergy and biomaterial feedstocks.
Identification of cell wall characteristics desirable for biorefining applications is crucial for lignocellulosic biomass improvement. However, the task of scoring biomass quality is often complicated by the lack of a reference for a given feedstock.
A multidimensional cell wall analysis was performed to generate a reference profile for leaf
and stem biomass from several miscanthus genotypes harvested at three developmentally distinct time points. A comprehensive suite of 155 monoclonal antibodies was used to monitor
changes in distribution, structure and extractability of noncellulosic cell wall matrix glycans.
Glycan microarrays complemented with immunohistochemistry elucidated the nature of
compositional variation, and in situ distribution of carbohydrate epitopes. Key observations
demonstrated that there are crucial differences in miscanthus cell wall glycomes, which may
impact biomass amenability to deconstruction.
For the first time, variations in miscanthus cell wall glycan components were comprehensively characterized across different harvests, organs and genotypes, to generate a representative reference profile for miscanthus cell wall biomass. Ultimately, this portrait of the
miscanthus cell wall will help to steer breeding and genetic engineering strategies for the
development of superior energy crops.
Introduction
Global hydrocarbon availability has reportedly been increased
thanks to modern hydraulic fracturing (Norris et al., 2016).
However, studies have reported that these extraction technologies
employ toxic, allergenic, mutagenic and carcinogenic chemical
additives, and cause the surfacing of radioactive materials (Lechtenb€
ohmer et al., 2011). At a time when sustainability is crucial,
‘greener’ solutions for the production of fuel and other bioproducts may be found in lignocellulosic biomass. Dedicated secondgeneration bioenergy crops can provide such biomass, and the
Miscanthus genus contains species with high potential as sustainable biomass providers (Carroll & Somerville, 2009).
Considering their high biomass yields, perenniality, C4 carbon
fixation, potential for soil carbon sequestration, reduced soil erosion and low fertilizer requirement (Clifton-Brown et al., 2013;
van der Weijde et al., 2013), the most relevant miscanthus varieties include M. sinensis, M. sacchariflorus and M. 9 giganteus
(Heaton et al., 2008; Dwiyanti et al., 2013; Liu et al., 2013). In
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the UK, miscanthus is planted during spring, and once established can be harvested annually for up to 15 yr (DEFRA, 2007).
New shoots typically emerge by midspring and grow rapidly in
the following months, reaching several metres in height by midsummer, depending on genotype. Autumn frosts trigger senescence, and senescing miscanthus remobilizes nutrients from
above-ground organs to rhizomes. For this reason, only completely senesced plants are usually harvested, thus ensuring crop
regrowth in the subsequent season.
Despite numerous potential biorefining applications, lignocellulosic biomass still remains largely untapped as a consequence of cell wall recalcitrance, which refers to the resistance
of cell walls to deconstruction (Himmel et al., 2007), dictated
by relative abundances and interactions between cell wall components (Pauly & Keegstra, 2010). Accordingly, for the aim of
efficient utilization of cell wall biomass as a renewable source
of useful molecules, it is vital to further our knowledge regarding how walls are assembled, in terms of both composition
and structure.
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distribution and reproduction in any medium, provided the original work is properly cited.
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Commelinoid monocots, which include grasses, contain cell
walls that are distinct from those found in other plant taxa. In
addition to lignin, these cell walls contain high percentages of cellulose (microfibrillar 1?4-b-glucan), low percentages of xyloglucan (XG; 1?4-b-glucan substituted by xylosyl residues), mixedlinkage 1?3, 1?4-b-glucan (MLG), and a high abundance of
1?4-b-xylan. Xylans frequently contain acetyl, arabinosyl and/
or glucuronyl substituents attached to some backbone xylose
residues (Carpita, 1996; Scheller & Ulvskov, 2010), hence the
designations, arabinoxylan (AX) and glucuronoarabinoxylan
(GAX). Grass cell walls also contain small amounts of pectins,
which are a-galacturonate-rich polysaccharides, and are thought
to consist essentially of three interconnected domains joined
together by glycosidic bonds: homogalacturonan (HG), rhamnogalacturonan-I (RG-I) and rhamnogalacturonan-II (RG-II)
(O’Neill et al., 1990; Fry, 2010; Atmodjo et al., 2013). HG
makes up most of cell wall pectin and comprises unbranched
chains of a-galacturonate residues, joined by 1?4-bonds, which
may be methyl-esterified (Atmodjo et al., 2013). Additionally,
other less abundant components may be found, such as structural
proteins and hydroxycinnamates (HCAs). The complexity of cell
wall composition is increased by the fact that cell wall components are interconnected via processes that are not yet completely
understood (McCann & Carpita, 2015; Park & Cosgrove,
2015), to form molecular structures that maintain the integrity
of plant tissues and resist exogenous attack (Pauly & Keegstra,
2008; Chowdhury et al., 2014). Consequently, understanding
these complex and intricate networks is a daunting task, especially if we consider that c. 10% of plant genomes is associated
with cell wall assembly and disassembly (Carpita & McCann,
2015).
Research aimed at improving lignocellulosic biomass for biorefining applications has motivated considerable advances in our
understanding of plant cell walls. Identification of desirable cell
wall characteristics, and the development of crops containing
such features, are crucial steps for lignocellulosic biorefining optimization. However, as all cell walls are not identically composed,
it is often difficult to systematically assess the quality of a given
feedstock in relation to others, particularly between different
studies. Here we present a reference cell wall profile for miscanthus leaf and stem biomass harvested at three developmentally distinct time points. This reference profile is particularly focused on
cell wall glycan components, given that glycans compose the bulk
of grass cell walls and possess the most wide-ranging potential for
industrial valorization (Lucia, 2008; Uraki & Koda, 2015).
A developmentally focused study of structural glycan composition and distribution in miscanthus organs and/or tissues is relevant not only for the optimization of lignocellulosic biomass
utilization as a feedstock for renewable bioproduct and bioenergy
solutions, but also to improve our fundamental understanding of
the cell wall. Previous work based on Fourier transform midinfrared spectroscopy (FTIR) studies suggested that structural
polysaccharides are major contributors to compositional variability during stem development and between organs (da Costa et al.,
2014). In the present study, we identify components responsible
for this compositional variability and discuss several relevant
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observations related to composition, structure and possible interactions between cell wall components. Moreover, a large and
diverse set of plant glycan-directed monoclonal antibodies
(mAbs) was used to monitor changes in matrix glycan distribution, structure and extractability. This suite of molecular probes
possesses enough variability to allow the identification of most
classes of noncellulosic plant cell wall polysaccharides (Pattathil
et al., 2010, 2015). For the first time, this array of glycan-directed
mAbs was used to comprehensively characterize variations in the
cell wall glycome of miscanthus across different harvests, organs
and genotypes, to generate a representative reference profile for
miscanthus cell wall biomass.
Several global breeding programmes focus on harnessing the
genotypic and phenotypic variation among and within miscanthus species with the aim of genetically improving miscanthus
traits relevant to the enhancement of biomass quality and yield
(Heaton et al., 2004; Yan et al., 2012; Robson et al., 2013; van
der Weijde et al., 2016). The detailed portrait of miscanthus cell
wall presented here will contribute to these efforts, as it provides
relevant information for the tailoring of miscanthus varieties with
preferable characteristics for conversion to biofuels and other
biomaterials.
Materials and Methods
Plant material
A general characterization of miscanthus cell wall has been
reported (da Costa et al., 2014). Here we present more detailed
studies based on a subset of the genotypes used in the original
study: M. 9 giganteus (gig01), M. sinensis (sin08, sin09, sin11,
sin13, sin15), M. sacchariflorus (sac01) and a nonspecified interspecific hybrid (hyb03). Experimental plots and growth conditions have been described previously (Allison et al., 2011; da
Costa et al., 2014).
Cell wall material
All compositional analyses and Clostridium phytofermentans
bioassays were performed on purified cell wall material (CWM).
Individual leaf and stem samples were collected from single tillers
harvested at time points corresponding to three developmental
stages: 10 wk after shoot emergence, when plants were actively
growing; 18 wk, when plant growth had reduced to a minimal
rate, peak biomass; 42 wk, senesced stage. CWM was prepared as
described in da Costa et al. (2015).
Neutral monosaccharides
Acid hydrolysis was performed as described in Sluiter et al.
(2012): 10 mg of CWM were weighed into 10 ml Pyrex glass
tubes and 100 ll H2SO4 (72% w/w) was added. Sealed tubes
were left at 30°C for 1 h. Samples were diluted to 4% H2SO4
(w/w) and autoclaved at 121°C for 1 h. Once at room temperature, tubes were centrifuged to produce a supernatant; 400 ll of
diluted (1 : 200) samples was transferred to filter vials (0.45 lm
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nylon filter; Thomson Instrument Company, Oceanside, CA,
USA). Carbohydrate separation was by high-performance anionexchange chromatography with pulsed amperometric detection
(HPAEC-PAD) using a gold working electrode and Ag/AgCl reference electrode. The ICS-5000 ion chromatography system
(Dionex, Sunnyvale, CA, USA) was operated at 45°C using a
CarboPac SA10 column with a CarboPac SA10G guard column.
An eluent generator prepared 0.001 M KOH for 14 min isocratic
elution at 1.5 ml min 1. Calibration standards were used for
monosaccharide identification and quantitation. In order to
resolve galactose (Gal) which coelutes with rhamnose at 45°C,
the HPAEC-PAD method was adapted by reducing the flow rate
to 1.2 ml min 1 and temperature to 30°C.
Acetyl esters
Cell wall-bound acetate was released by an alkaline saponification
procedure modified from Manabe et al. (2011). CWM (10 mg)
was incubated in 500 ll 0.1 M KOH for 16 h with constant
shaking (21°C/150 rpm). Samples were centrifuged at 2500 g for
5 min, 100 ll of the supernatants were mixed with 900 ll
0.005 M H2SO4 containing 0.005 M crotonic acid as an internal
standard. The mixtures were filtered through 0.45 lm syringe filters (Millipore Corporation, Billerica, MA, USA) and 25 ll analysed on a high-performance liquid chromatography (HPLC)
system fitted with a refractive-index detector (Jasco, Great Dunmow, Essex, UK), equipped with a Rezex ROA-organic acid H+
column (Phenomenex, Torrance, CA, USA), kept at 35°C, with
a 0.005 M H2SO4 mobile phase flowing at 0.6 ml min 1 for
16 min. Supernatant acetate concentrations were determined
using a concentration gradient of an acetic acid standard.
Hydroxycinnamoyl esters
Ester-linked HCAs were released using an alkaline saponification
method adapted from Buanafina et al. (2006). CWM (10 mg)
was mixed with 5 ml 1 M KOH under a flow of N2 to reduce
sample oxidation, followed by dark incubation for 16 h with constant shaking (21°C/150 rpm). Samples were centrifuged at
2500 g for 5 min, extracts were transferred to new tubes and pellets were washed with 4 ml methanol (100%). Wash supernatants
were combined with the extracts and solubilized carbohydrate
was precipitated at 80°C/20 min. After centrifugation (2500 g;
5 min) supernatants were collected in new tubes, pellets were
washed with 1 ml methanol (100%) and all supernatants were
combined. After methanol evaporation and sample acidification,
HCAs were recovered by reverse-phase C18 solid-phase extraction
(Sep-Pak C18 Vac RC cartridges, 500 mg, 3 cm3, 55–105 lm;
Waters Corporation, Milford, MA, USA). The resulting samples
were dried under N2 and reconstituted in 200 ll methanol (70%
v/v); 20 ll was injected for analysis on a reverse-phase HPLC system with a diode array detector (RP-HPLC-DAD; Waters Corp.)
set to collect UV/visible spectra at 240–400 nm. A radial compression column was used (Nova-Pak C18 Radial-Pak Cartridge,
4 lm particle size; Waters Corp.), with methanol (100%) and
acetic acid (5% v/v) as eluents in a 25 min linear 20–70%
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methanol gradient elution at 2 ml min 1. Identity and concentration of monomeric ferulic acid and p-coumaric acid were determined using authentic standards.
Statistical analysis
All descriptive statistics, ANOVA, Tukey’s tests and correlations
were calculated using Statistica (v. 8.0; StatSoft, Tulsa, Oklahoma) at a 5% significance level (a = 0.05). Effect sizes were calculated as g2 statistics (Cohen, 1973; Levine & Hullett, 2002):
g2 = SSeffect/SStotal (where SS is the sum of squares).
Glycome profiling
Sequential extractions of CWM, phenol-sulphuric acid total
carbohydrate estimation assays and enzyme-linked immunosorbent assays of resultant extracts, all part of the glycome profiling study, were carried out as described previously (DeMartini
et al., 2011; Pattathil et al., 2012). Sequential extractions consisted of using increasingly harsher chemicals : ammonium
oxalate (0.05 M), sodium carbonate (0.05 M), 1 M KOH, 4 M
KOH, acidic sodium chlorite and 4 M KOH PC (postchlorite
treatment). The sequential extracts were screened with a comprehensive suite of noncellulosic glycan epitope-directed mAbs
(Pattathil et al., 2010) obtained from the Complex Carbohydrate Research Center laboratory stocks (CCRC, JIM and
MAC series) or from BioSupplies (Bundoora, Vic., Australia)
(BG1, LAMP). Supporting Information Table S1 describes all
mAbs used. Heat maps representing glycome profiling data
were produced as previously described (Pattathil et al., 2012).
Principal components analysis (PCA) of glycome profiling data
was performed using the Eigenvector PLS Toolbox (v. 7.0.3;
Eigenvector Research, Wenatchee, WA, USA) within MatLab
(v. R2010b; MathWorks, Natick, MA, USA).
Lignin measurement
Lignin content of the residues resulting from the sequential
extraction was determined using the acetyl bromide procedure, as
previously described (da Costa et al., 2014).
C. phytofermentans bioassay of biomass digestibility
A previously described bioassay (Lee et al., 2012a,b) which uses
Clostridium phytofermentans-mediated ethanol fermentation
yields as digestibility indicators was used to estimate CWM
amenability to deconstruction.
In situ immunolabelling
Sections from the middle portions of leaf blades and stem internodes, both located halfway through the length of the tiller, from
gig01 (M. 9 giganteus) plants were sampled at peak biomass and
immersed in a fixative solution consisting of paraformaldehyde
(1.6% v/v) with glutaraldehyde (0.2% v/v) in 0.025M sodium
phosphate buffer (pH = 7.1).
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In situ cell wall glycan epitopes were detected by fluorescence
immunolabelling according to procedures described elsewhere
(Avci et al., 2012). Briefly, samples were dehydrated using a
graded ethanol series, infiltrated and embedded in LR White
resin (Ted Pella, Redding, CA, USA) in gelatine capsules. Resin
was polymerized at 4°C under UV light (365 nm). Transverse
semithin sections (250 nm) were cut with a Leica EM UC6 ultramicrotome (Leica Microsystems, Buffalo Grove, IL, USA). Sections were blocked with 3% (w/v) nonfat dry milk in 0.01 M
potassium phosphate-buffered saline containing 0.5 M NaCl
(KPBS; pH = 7.1) for 30 min and washed with the same buffer.
Undiluted hybridoma supernatants containing the mAbs were
applied to the sections, incubated for 90 min and then washed
with KPBS. Goat anti-mouse immunoglobulin-G (IgG) or antirat IgG conjugated to Alexa-fluor 488 (Invitrogen, Waltham,
MA, USA) diluted 1 : 100 in KPBS was applied. After 90 min,
sections were washed with KPBS and deionized H2O. For certain
mAbs known to only bind to de-esterified forms of the epitopes,
sections were subjected to a base treatment with 0.1 M KOH (1 h
followed by three washes with deionized H2O), before blocking
and mAb application. Microscopic inspection was performed
using an Eclipse 80i microscope (Nikon, Melville, NY, USA)
equipped with epifluorescence optics. Images were captured with
a Nikon DS-Ri1 camera head using NIS-Elements Basic
Research software (Nikon).
Results
Relative abundances of cell wall monosaccharides vary
significantly in stem and leaf biomass throughout plant
growth
Plant cell wall glycans and their monosaccharide constituents represent valuable biorefinery feedstocks. Monosaccharide compositions were determined for each leaf and stem sample from the
eight genotypes examined in this study, harvested at three developmental stages: active growth, peak biomass and senescence
(Table 1). Further information concerning the distribution of
monosaccharide content measurements across eight genotypes is
provided in Figs S1 and S2.
Fucose (Fuc) and Gal together represented < 2% of miscanthus CWM dry weight (DW). Glucose (Glc) was the most abundant monosaccharide in miscanthus CWM, followed by xylose
(Xyl), comprising, on average, 44.2% and 14.9% DW of all samples analysed, respectively (Table 1). Overall, Glc and Xyl contents were significantly higher in stems than in leaves (P < 0.001).
However, Xyl content did not differ significantly (P = 0.880)
between senesced leaves and stems (Table 1). Glc abundance differed significantly (P < 0.001) between harvests, as values were
higher earlier in development than at senescence. Xyl content also
varied significantly (P < 0.001) throughout development,
although differently for each organ. In common with Glc trends,
the abundance of Xyl in stem cell walls was higher at the first harvest than at senescence. However, in leaves, Xyl content was
higher in senesced material than at active growth.
Mean arabinose (Ara) content was c. 2% of CWM DW
(Table 1). Ara abundance primarily varied according to organ
origin (P < 0.001; g2 = 0.6966), with leaves containing almost
twice the Ara content of stems. Additionally, Ara abundance varied significantly (P < 0.001) between harvests, decreasing in consecutive harvests. Although Ara and Xyl may originate from
different classes of cell wall polysaccharides, in grasses like miscanthus these monosaccharides should derive primarily from AX
(Carpita, 1996). Xylan arabinosylation may be estimated by calculating Ara/Xyl ratios (Rancour et al., 2012). Organ origin was
the main source of variation in Ara/Xyl ratios (P < 0.001;
g2 = 0.6841), as values were higher in leaves than in stems by a
factor of 2.33 at active growth, 2.11 at peak biomass and 1.78 at
senescence (Table 1). In leaves, Ara/Xyl ratios suggested altered
arabinosylation degrees as plants matured, with a decline from
Table 1 Composition of miscanthus biomass expressed as percentage of cell wall material (CWM) DW (%CWM)
Active growth
Total sugar
Glucose
Xylose
Arabinose
Galactose
Fucose
Ara/Xyl
Lignin
Acetate
p-Coumaric acid
Ferulic acid
Released ethanol
Peak biomass
Senescence
Leaf
Stem
Leaf
Stem
Leaf
Stem
61.7 9.9
43.6 6.6
14.2 2.8
2.89 0.63
0.84 0.18
0.20 0.05
0.21 0.04
18.1 0.9
3.15 0.32
0.75 0.29
0.43 0.17
51.8 2.5
66.5 9.2
48.4 7.2
16.2 2.5
1.43 0.32
0.29 0.12
0.22 0.06
0.09 0.02
19.1 1.8
4.77 0.36
1.18 0.63
0.41 0.15
49.4 3.9
57.3 5.3
40.4 3.8
13.3 1.7
2.55 0.51
0.88 0.20
0.23 0.05
0.19 0.04
19.5 0.6
3.36 0.37
0.66 0.21
0.32 0.11
48.4 2.5
63.2 6.3
46.1 4.5
15.2 2.4
1.36 0.33
0.33 0.11
0.21 0.06
0.09 0.02
21.8 1.6
4.70 0.38
1.14 0.64
0.35 0.14
46.2 2.3
60.6 5.4
41.8 3.2
15.4 2.2
2.44 0.35
0.77 0.17
0.22 0.07
0.16 0.03
22.1 1.5
4.19 0.71
0.72 0.25
0.30 0.13
39.4 1.4
62.2 7.3
45.0 5.3
15.4 2.2
1.32 0.20
0.34 0.11
0.20 0.07
0.09 0.02
23.8 0.7
4.66 0.53
1.10 0.45
0.32 0.12
40.1 1.4
Values are means SD of leaf or stem biomass from eight genotypes at each individual developmental stage. Total sugar values represent the mean of the
sum of all quantified monosaccharides in each CWM sample collected from leaf or stem at each developmental stage. Ara/Xyl is the mean ratio of
arabinose : xylose in the samples. Lignin values were taken from da Costa et al. (2014) and means recalculated for the eight genotypes used in the present
study, instead of the original 25. Released ethanol values are expressed as mg ethanol g–1 dry CWM and consist of supernatant ethanol concentrations
after 72 h incubation with Clostridium phytofermentans.
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active growth to peak biomass ( 9.5%) and between peak
biomass and senescence ( 15.8%). By contrast, in stems the
ratios did not vary substantially between harvests (Fig. S3).
Cell wall acetylation increases in leaf biomass as plants
mature, but remains constant in stems
The biological role of cell wall polymer acetylation is poorly
understood (Manabe et al., 2011, 2013), but may be involved in
xylan–cellulose interactions (Busse-Wicher et al., 2014), which
influence recalcitrance and, subsequently, biomass conversion.
Acetate release was measured by 0.1 M KOH-mediated saponification; that is, acetyl ester linkage cleavage (Marcus et al., 2010;
J€onsson et al., 2013). Significant (P < 0.001) differences between
organs were observed in released acetate abundance. However,
the degree of cell wall acetylation did not vary equally in each
organ throughout development. Acetate content of leaves differed
significantly between developmental stages (P < 0.001), as mean
acetylation degree increased in consecutive harvests. By contrast,
in the CWM of stems, measured acetate did not change significantly throughout plant development (P = 0.525), with an average value of 4.7% DW at all developmental stages (Table 1;
Fig. S4).
More FA and pCA are observed in younger plant cell walls
Grass cell wall polymers are abundantly feruloylated and
p-coumaroylated (Carpita, 1996). Ferulic acid (FA), which is predominantly ester-linked to AX and GAX, has been associated
with digestibility, while p-coumaric acid (pCA) appears mostly
esterified to lignin (Lu & Ralph, 1999). Ester-linked hydroxycinnamoyl quantitation was performed after cell wall saponification
with 1 M KOH. FA content was not significantly different
between leaf and stem (P = 0.153), but at active growth, mean
FA content was higher than in later harvests by at least 34% in
leaves, and 17% in stems (Table 1; Fig. S5). Conversely, pCA
release was significantly different between organs (P < 0.001),
with mean pCA content in stems being 50% higher than in leaves
(Table 1).
A reference cell wall glycome profile allows identification of
key glycan modifications in the cell wall of developing
plants
Changes in matrix glycan composition, structure and extractability were monitored in miscanthus CWM using glycome profiling, which employs increasingly harsh, sequential multistep
extractions to fractionate the cell wall (Pattathil et al., 2012; Li
et al., 2014). Most noncellulosic glycans were extractable before
biomass delignification with acidic sodium chlorite, and the 1 M
KOH fractions contained the largest portion of extracted glycans.
Total released carbohydrate present in each extract and the composition of the residues remaining after cell wall fractionation are
shown in Fig. S6 and Tables S2, S3.
Glycome profiling data are summarized as heat maps,
where the mean (Fig. 1), and the standard deviation
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(Fig. S7) of binding intensities for each glycan-directed mAb
across all miscanthus genotypes are depicted for each harvest
and organ.
Antibodies directed at XG epitopes showed highest binding
signals in the 4 M KOH extracts (both pre- and post-sodium
chlorite treatment) with more diverse and higher abundance of
XG epitopes being removed from stems than from leaves, particularly before delignification. This is exemplified by the significantly increased abundance of XG epitopes recognized by
nonfucosylated (nonfuc) XG-4, nonfuc XG-5 and xylan-1/XG
groups of mAbs. Among the XG epitopes extracted with 1 M
KOH, those detected by the nonfuc XG-1 and nonfuc XG-2
groups of mAbs were more abundant in active growth leaf samples than in mature leaves, or in stems.
Xylan epitopes were found in all extracts, but their presence
and abundance varied in different organs and harvests. For the
most weakly integrated xylans present in oxalate extracts, epitopes bound by the xylan-4 and xylan-7 mAb subclasses were
present in leaf and stem, but marginally more abundant in
actively growing stems. High abundance of xylan epitopes was
apparent in most carbonate extracts, with the exception of
leaves from peak biomass and senesced harvests. In these cases,
there were lower signals for xylan-6 mAbs. In the case of more
tightly bound xylans, released with 1 M and 4 M KOH (both
pre- and postchlorite), epitopes recognized by the xylan-4
through xylan-7 groups of mAbs were highly abundant. In
general, no substantial variations in the abundance of these
epitopes were noted between organs or developmental stages.
Interestingly in chlorite extracts, a reduced abundance of xylan
epitopes that are specifically recognized by the xylan-6 groups
of mAbs was detected.
Binding intensities of mAbs directed at 1?3-b-glucan (callose) and MLG increased with harsher extractions and were more
abundant in leaves. Exceptions are the 4 M KOH extracts, where
binding was similar between organs.
Homogalacturonan backbone-1 epitopes occurred in all fractions with few developmental and organ differences, except for
slightly higher binding intensities in the oxalate and 1 M
KOH extracts from leaves. RG-I backbone epitope detection
was low, almost exclusively perceptible in strong alkaline and
chlorite extracts, but subtly more abundant in leaf extracts.
The RG-I/AG class of mAbs, which detects pectic arabinogalactan epitopes, in general showed higher abundance in
oxalate, carbonate and 1 M KOH extracts from active growth
and, overall, leaf extracts exhibited marginally increased abundance of these epitopes compared with stem samples. A higher
abundance of these epitopes in leaves from all developmental
stages is also exhibited in 4 M KOH PC extracts. Glycan epitopes bound by the AG-1 to AG-4 groups of mAbs may occur
in arabinogalactan-containing polysaccharides as well as in arabinogalactan proteins (AGPs). For these mAbs, the highest signals were detected in oxalate extracts from actively growing
plants, mainly in stems. In chlorite extracts, especially from
stems, binding signals for some AG-4 mAbs were increased,
suggesting that a proportion of these epitopes could not be
released before delignification.
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Fig. 1 Glycome profile of miscanthus cell wall material (CWM) extracted sequentially with ammonium oxalate, sodium carbonate, 1 M KOH, 4 M KOH,
sodium chlorite and 4 M KOH postchlorite treatment (PC) as explained in the Materials and Methods section. Corresponding organs and developmental stages
are labelled below each profile: AGr, actively growing; PBm, peak biomass; SSn, senesced. Each extract was probed against an array of plant glycan-directed
monoclonal antibodies (Supporting Information Table S1 for a list of all mAbs). Antibody binding strength based on optical density (OD) is presented as a
colour gradient ranging from black (no binding) and red (intermediate binding), to yellow (strongest binding) (OD = 1.3). Binding intensities are the averages of
all genotypes studied here. Red bars on the top indicate the average amount of sugars recovered in the solubilized extracts g–1 CWM (mg g 1).
In situ immunolabelling revealed organ-specific distribution
patterns of cell wall glycan epitopes
Immunohistochemistry of M. 9 giganteus stem and leaf midrib
sections using key mAbs recognizing distinct glycan epitopes
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(Table 2) revealed epitope distributions at the cellular level, thus
verifying glycome profiling results in situ.
Nonfucosylated XG epitopes probed with CCRC-M88
revealed strong labelling in phloem but fainter labelling in xylem
of leaves and stems (Fig. 2). However, fluorescence patterns
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Table 2 Cell wall glycan-directed monoclonal antibodies (mAbs) used in
the study of in situ immunolabelling of Miscanthus 9 giganteus leaf and
stem tissues (further information on all mAbs used here can be found in
Supporting Information Table S1)
mAb
mAb subclass – based on Pattathil et al. (2010)
CCRC-M88
CCRC-M1
CCRC-M154
CCRC-M155BT
CCRC-M149
BG1
CCRC-M38
CCRC-M128
Nonfucosylated xyloglucan-2
Fucosylated xyloglucan
Xylan-4 (arabinosylated xylan)
Xylan-5 (Me-GlcA substituted xylan)
Xylan-7 (xylan backbone DP≥4)
Mixed-linkage (1?3, 1?4)-b-glucan
Demethyl-esterified homogalacturonan backbone-1
RG-I/AG (arabinogalactan side chains of
rhamnogalacturonan-I)
BT
Used in combination with a 0.1 M KOH base treatment of the sections
before immunolabelling.
differed between organs, as labelling of parenchymatous and sclerenchymatous structures was more perceptible in stems. Probing
of fucosylated XG with CCRC-M1 showed more abundant
labelling in stem cell walls, but mostly restricted to phloem.
Xylan-directed mAbs CCRC-M154 (xylan-4), CCRC-M149
(xylan-7) and CCRC-M155 (xylan-5) displayed varying
labelling patterns in different organs (Fig. 2). With CCRCM154, which is selective for arabinose-substituted xylans (Schmidt et al., 2015), labelling was low in stems, but was
detected in most leaf cell walls, although more intensely in
the phloem. CCRC-M149, which binds to the xylan backbone, abundantly labelled cell walls of most cell types, but
less in leaf mesophyll and interfascicular parenchyma.
Labelling of epidermal and sclerenchymatous thickenings was
particularly visible. Very low reactivity was detected initially
with CCRC-M155, which binds to a 4-O-Me GlcAsubstituted xylan epitope (M. G. Hahn et al., unpublished)
(Fig. S8). All of the xylan-directed mAbs used in this study
were generated using alkali-extracted de-esterified xylans and
therefore may not recognize acetylated xylan epitopes in vivo
(Li et al., 2014). To test for this, 0.1 M KOH was employed
to remove ester-linked groups in the organ sections, thus rendering xylan epitopes accessible. Epitope recognition by
CCRC-M155 became abundant after this base treatment
(Fig. 2), revealing where at least some esterified xylan epitopes
occur in intact cell walls. Labelling was predominant in
phloem and xylem parenchyma, although more widespread
patterns were apparent in leaves, including walls of bundle
sheath, epidermal and stomatal subsidiary cells. Additionally,
labelling was more intense in portions of the outer layers of
the walls in foliar parenchymatous ground tissue.
In accordance with glycome profiling results, labelling with
CCRC-M174 (galactomannan-directed) was negligible in analysed sections (Fig. S8).
The BG1 mAb, which binds MLG epitopes (Meikle et al.,
1994), showed more intense labelling patterns in leaf sections
(Fig. 2), where fluorescence was visible in most cell types, except
in walls of metaxylem, mesophyll and stomatal subsidiary cells.
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Labelling of AG side chains of RG-I with CCRC-M128 was
concentrated in membranous structures. De-methyl-esterified
HG epitope distributions probed with CCRC-M38 were similar
in the vascular bundles of stems and leaves, as only phloem and
protoxylem were labelled (Fig. 2). However, patterns differed
between organs, as leaf parenchyma and mesophyll showed
higher accumulation of CCRC-M38 epitopes, particularly on
middle lamellae and lining intercellular spaces of cell junctions.
Miscanthus biomass amenability to conversion depends on
organ and harvest
A C. phytofermentans bioassay was employed to estimate how miscanthus CWM recalcitrance varied between leaf and stem samples and throughout plant maturation. Mean released ethanol
yields, expressed as mg ethanol g–1 cell wall DW, ranged from
40.07 mg g 1 in senesced stem to 51.83 mg g 1 in active growth
leaf. Ethanol yield decreased as plants matured and was higher in
leaves than in stems, except for senesced samples, where the
values did not vary substantially (Table 1).
Discussion
Miscanthus cell wall glycans
Structural polysaccharides are major contributors to compositional differentiation between miscanthus organs and harvests (da
Costa et al., 2014). Here, we explored these differences further
and identified typical cell wall characteristics of miscanthus
organs from different harvests (summarized in Fig. 3).
In Poales cell walls, Glc is primarily derived from cellulose, but
some Glc is also found in MLG and XG, which are in much
lower abundance in Poales walls (Carpita, 1996). The higher Glc
content in stems than in leaves, and in younger plants than at
senescence, shown in this study have also been reported elsewhere
for miscanthus (Le Ngoc Huyen et al., 2010; van der Weijde
et al., 2016) and for Brachypodium distachyon (Rancour et al.,
2012). Given the supportive role of stems, higher total sugar content implies more abundant structural polysaccharides, which
make up the major load-bearing network in plant cell walls
(Harris & Stone, 2008; Xu, 2010; Leroux, 2012).
The combined Ara and Xyl contents suggest that the abundance of AX, which is the principal glycan from the Poales walls
from which these monosaccharides arise, does not vary considerably between organs. However, Ara content and Ara/Xyl ratios
were approximately twofold higher in leaves, suggesting higher
arabinosylation than in stems (Table 1; Fig. S3). This agrees with
higher AX branching reported in Saccharum officinarum leaves
than in culms (de Souza et al., 2013). It has also been shown that
highly substituted AX is prominent in primary cell walls, while
less substituted xylans are associated with secondary cell wall and
lignification (Suzuki et al., 2000).
In grasses, pectins and AGPs are the main Gal-containing cell
wall polysaccharides (Carpita, 1996; Ishii, 1997; O’Neill & York,
2003). Higher Gal in foliar biomass may thus suggest more
abundant pectin and/or AGPs in leaves than in stems.
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Fig. 2 Immunofluorescent labelling of cell wall glycan epitopes in transverse sections from leaves and stems from Micanthus 9 giganteus (gig01).
Immunolabelling studies were preceded by microscopic inspection of sections stained with toluidine blue to characterize their histological complexity. Used
mAbs: CCRC-M88, nonfucosylated xyloglucan (XG); CCRC-M1, fucosylated XG; CCRC-M154, Ara-substituted xylan (Xylan-4); CCRC-M149, xylan
backbone (Xylan-7); CCRC-M155, Me-GlcA-substituted xylan (Xylan-5 de-esterified epitopes); BG1, mixed-linkage (1?3, 1?4)-b-glucan; CCRC-M128,
arabinogalactan side chains of rhamnogalacturonan-I (RG-I/AG); CCRC-M38, demethyl-esterified homogalacturonan (HG-backbone-1). More details
regarding the mAbs are available in Supporting Information Table S1. For CCRC-M155, results are shown for immunolabelling of sections after a base
treatment (BT) with 0.1 M KOH, which removed ester-linked substituents from the walls (nonbase-treated control for CCRC-M155 may be found in
Fig. S8). abe, abaxial surface epidermis; bs, bundle sheath; e, epidermis; gt, parenchymatous ground tissue; is, intercellular space; mf, mesophyll cells; mx,
metaxylem; px, protoxylem; sf, sclerenchyma fibres; st, stomatal complex; xp, xylem parenchyma. Bars, 100 lm.
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glycome are highlighted in standard deviation heat maps
(Fig. S7). To clarify the significance of these heterogeneities
and determine glycan classes that contribute to the distinctiveness between organs and harvests, PCA was performed as a
discriminant tool.
Increasingly harsher extractants produce qualitatively distinct
cell wall fractions, demonstrating that different polysaccharide
classes are not equally integrated into the overall cell wall structure The various cell wall extracts formed clusters along principal component one (PC1; Fig. 5a). PC1 loadings (Fig. S10)
indicated that clusters on the negative side of PC1 (oxalate and
carbonate) correspond to extracts with typically stronger binding
by pectic epitope-directed mAbs. Conversely, clusters on the positive side of PC1 (4 M KOH and 4 M KOH PC) showed higher
binding by XG- and xylan-directed mAbs. These results suggest
that there are differences between the walls in leaf vs stem cells in
terms of how firmly different polysaccharide types are integrated
into the wall matrix. In chlorite extracts, xylan-directed mAbs
showed higher signals, but MLG and various pectin epitopes
were also detected. MLG binding signals were detected in all 1 M
KOH extracts, but were more abundant in actively growing
plants. This agrees with findings that, despite higher MLG accumulation in younger plants, it is also found in mature organs
(Vega-Sanchez et al., 2013). As MLG is released using extractants
of different stringency, varying solubility may depend on ratios
between trisaccharide/tetrasaccharide MLG domains (Collins
et al., 2010; Vega-Sanchez et al., 2015).
Fig. 3 Mean percentage (%) composition of miscanthus cell wall biomass
DW. Overall indicates the composition for leaf and stem at all
developmental stages combined. Lignin values are reproduced from da
Costa et al. (2014). Undetermined components may include uronic acids,
ferulate dimers and oligomers, methyl groups, proteins and minerals. FA,
ester-linked ferulic acid; pCA, ester-linked p-coumaric acid; AGr, active
growth; PBm, peak biomass; SSn, senescence; L, leaf; S, stem.
Key glycome modifications in miscanthus cell walls
A miscanthus reference glycome profile (Fig. 1) represents a
powerful tool for future studies, allowing comparisons with
other genotypes or lignocellulosic feedstocks. To facilitate
comparisons between extracts, organs and harvests, this reference profile was graphically reinterpreted (Figs 4, S9). Furthermore, the most heterogeneous portions of the cell wall
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Miscanthus leaves and stems present distinct matrix cell wall
glycomes and hence have different overall wall structures For
each sequential extraction step, a PCA model revealed two separate clusters: one comprising glycome profiles from stems, and
another from leaves (Figs 5b–e, S11). Depending on the PC
responsible for the separation, corresponding loadings identified
principal glycan contributors to organ-dependent variation. For
leaf oxalate and carbonate extracts, higher RG-I/AG epitope
binding is a main diverging factor in relation to stems. In carbonate extracts, binding to easily extracted xylan epitopes is also a
discriminant feature between organs, as stem samples exhibit
stronger binding by xylan-5, xylan-6 and xylan-7 groups of
mAbs. In 1 M KOH extracts, organ differentiation primarily
derives from higher signals in leaves for most glycan epitopes,
particularly for pectins and b-glucans. For 4 M KOH extracts
(pre- and postchlorite), stems typically showed higher binding
for XG and xylan-5, xylan-6 and xylan-7 mAbs. In leaves, 4 M
KOH extracts showed higher binding for most xylan-3, xylan-4
and RG-I mAbs.
Stems and leaves are differently modified throughout development Individual organ models revealed no PCA clusters for any
of the stem extracts nor for the three harshest leaf cell wall
extracts (Fig. S12), implying that epitope abundance variation
for these samples is not significant between harvests. However, in
oxalate, carbonate and 1 M KOH leaf extracts, differences in
noncellulosic glycan epitope abundances were sufficient to create
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Fig. 4 Mean binding values to different
classes of cell wall glycan epitopes released at
sequential extraction steps from leaf and
stem samples from miscanthus biomass at
three developmental stages (cf. Fig. 1 and
Supporting Information Fig. S9).
two clusters, one comprising peak biomass and senesced samples
(overlapped), and another from active growth (Fig. 5f–h).
Principal components analysis of glycome profiling data highlighted significant variation between harvests and organs. These
differences agree with FTIR-based predictions that structural
polysaccharides are major contributors to developmental and
organ-dependent compositional variation in miscanthus lignocellulosic biomass (da Costa et al., 2014). Considering both studies,
it appears that cell wall compositional variation between mature
and immature plants is primarily associated with nonmatrix cell
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wall components in stems, namely cellulose and lignin (da Costa
et al., 2014). In leaves, overall carbohydrate composition varies
less than in stems, but significant differences are seen in more
easily extractable and less abundant noncellulosic glycan
epitopes.
Tightly bound glycan epitopes and biomass recalcitrance Glycome profiling suggests that certain noncellulosic polysaccharide
components (namely, subclasses of XGs, heteroxylans, MLG and
pectins) are more firmly integrated in the cell wall matrix than
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Fig. 5 Principal components analysis of
glycome profiling data. Plot of principal
component 1 (PC1) and principal component
2 (PC2) scores for all samples (a; open
triangles in the 1 M KOH extract are stem
samples, and blue-filled triangles are leaf
samples), for samples extracted with 1 M
KOH (b), 4 M KOH (c), sodium chlorite (d)
and 4 M KOH postchlorite (PC) treatment
(e); (f, g, h) leaf samples extracted with
ammonium oxalate (f), sodium carbonate (g)
and 1 M KOH (h). Ellipses are intended to
highlight the structure of the data points.
AGr, active growth; PBm, peak biomass; SSn,
senesced stage.
others. Knowledge of the structural associations of these populations of glycans with other cell wall constituents is important to
improve our understanding of structural features underpinning
cell wall recalcitrance.
As sodium chlorite and 4 M KOH PC extracts contain glycans
thought to be either directly or indirectly associated with lignin
in the cell wall matrix, glycan epitopes detected in these extracts
are particularly informative with regard to cell wall recalcitrance.
Generally, signals in these extracts did not change notably during
development, but binding to most glycan epitopes was higher in
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chlorite extracts from leaf material than from stems (Figs 4, S11).
Most obviously for mAbs in the xylan-4, xylan-5, xylan-7, MLG
and several pectin-directed groups, the glycome profiling data
suggest that delignification generally removes more of these glycan epitopes from leaf cell walls than from stem. HG epitopes
released with sodium chlorite and 4 M KOH PC (Fig. 4) suggest
a pectin–lignin association, corroborated by the higher signals for
some RG-I and RG-1/AG-related mAbs in these extracts.
Pectin–lignin associations have been suggested for several species:
M. sinensis (de Souza et al., 2015), Populus trichocarpa
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(DeMartini et al., 2011), Panicum virgatum (Shen et al., 2013)
and Medicago sativa (Wi et al., 2005). Abundant MLG epitopes
in 4 M KOH PC extracts suggest that all harvests contain proportions of tightly bound MLG, only released after delignification. High MLG detection in harsher extracts was also observed
in glycome profiles of switchgrass (Shen et al., 2013), sugarcane
(de Souza et al., 2013) and corn stover (Li et al., 2014), and may
derive from tight MLG–cellulose associations (Carpita, 1996;
Fry, 2010; Kiemle et al., 2014). XG epitopes, particularly nonfuc
XG-1, nonfuc XG-2 and nonfuc XG-3 groups of mAbs, were
prominent in postchlorite extracts, suggesting that despite their
reduced abundance in grasses, structural features associated with
XG may inhibit sugar release from the cell wall matrix. Considering all extracts, and irrespective of organ origin or developmental
stage, signals for the xylan-4 to xylan-7 groups of mAbs (which
recognize diverse xylan epitopes) were most prominent in
postchlorite extracts. Hence, heteroxylans are likely to play
important roles in biomass recalcitrance, perhaps as a result of
the formation of ferulate–polysaccharide–lignin complexes that
crosslink the cell wall (Buanafina, 2009), and/or interactions
between arabinoxylan and cellulose.
Key dissimilarities between organs detected via glycome
profiling were verified by immunohistochemistry
In situ immunolabelling was performed using mAbs aimed at epitopes from the main noncellulosic miscanthus cell wall glycan
classes, and with distinct extractabilities, as revealed by glycome
profiling. The resulting labelling patterns suggest that distinct
glycan distributions are associated with organ and tissue structural requirements, which may influence biomass recalcitrance to
deconstruction.
More broadly disseminated XG epitopes in stem sections compared with leaves may derive from the higher abundance of these
glycans in stems, as observed in glycome profiling. As XG is
strongly wall-bound and presumed to interact with cellulose
(Hayashi, 1989), XG may be more relevant for stem cohesiveness
than for leaves.
Xylan-directed mAbs showed distinct labelling patterns, suggesting that M. 9 giganteus cell wall xylan abundance and structure vary depending on cell type. Labelling of primary walls of
M. 9 giganteus sections (Fig. 2) with CCRC-M154, which binds
arabinosylated xylans (Schmidt et al., 2015), is consistent with
reports that GAX occurs in interstitial spaces between primary
cell wall cellulose microfibrils (McCann & Carpita, 2008; Li
et al., 2014). When used with a base treatment, which removes
ester-linked substituents, epitopes recognized by CCRC-M155
are localized, in particular, to the phloem. By contrast, fewer of
the xylan backbone epitopes detected by CCRC-M149 are present in the phloem, but more are present in other cell walls, particularly where sclerified. Overall, these results suggest that
phloem xylans are more substituted than those in sclerenchyma.
In M. lutarioriparius (Cao et al., 2014), sugarcane (de Souza
et al., 2013) and maize (Suzuki et al., 2000), phloem is less lignified than sclerenchyma. Consequently, lignified cell walls may
contain less substituted xylans, while more substituted polymers
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coincide with lower lignification. Indeed, low-branched xylans
occur in all lignified walls, while most highly substituted xylans
occur in unlignified tissues (Suzuki et al., 2000). As lignin–carbohydrate associations affect cell wall recalcitrance to deconstruction, xylan substitution and localization pattern might influence
lignocellulose biorefining.
Reported MLG distribution patterns (Trethewey & Harris,
2002; Vega-Sanchez et al., 2012; Xue et al., 2013) are consistent
with BG1 labelling of sclerenchyma, phloem and protoxylem
cell walls (Fig. 2). To withstand the pressure and tension
required for plant support and vascular functions, the cell walls
of these organs must be structurally robust. MLG in sclerenchyma, phloem and protoxylem may indeed be contained in
the tightly bound MLG epitopes detected by glycome profiling
in the KOH and sodium chlorite extracts (Fig. 4), and contribute to structural cohesiveness. Similarly to MLG, leaf sections showed more abundant demethyl-esterified HG labelling
with CCRC-M38 (Fig. 2). HG demethyl-esterification promotes the formation of rigid pectin structures, enhancing cell
adhesion (Zhang & Staehelin, 1992; Mohnen, 1999; Anthon
& Barrett, 2006; Lionetti et al., 2010). Furthermore, HG
demethyl-esterification may increase in response to cellulose
depletion (Burton et al., 2000; Manfield et al., 2004; Wolf
et al., 2009). Given potential cell wall assembly roles, it is pertinent to clarify if recalcitrant MLG and demethyl-esterified HG
in leaves are adaptations for increased cell wall cohesiveness in
organs with lower cellulose content.
Biomass recalcitrance may be affected by several cell wall
features beyond lignin content
We have previously reported that the degree of miscanthus
biomass lignification does not completely account for biomass
digestibility (da Costa et al., 2014). In the present study, we
expand this conclusion and reveal that several cell wall features
may affect recalcitrance, particularly in leaf biomass, as evidenced
by correlations with C. phytofermentans fermentation yields
(Table S4).
Acetylation, in particular, shows strong negative relationships
with ethanol yields from leaves (r = 0.52; P < 0.01), suggesting
that acetylation is associated with recalcitrance-enhancing cell
wall features. Saccharification and fermentation limitation may
originate from acetyl-mediated steric hindrance to binding of
hydrolytic enzymes (Biely, 2012; Pawar et al., 2013) and/or
microbial inhibition (Gille & Pauly, 2012). Furthermore, acetylation degrees may affect secondary cell wall xylan–cellulose
interactions (Busse-Wicher et al., 2014) and influence plant
development, as supported by stunted growth phenotypes in
Arabidopsis mutants with reduced wall acetylation (Manabe et al.,
2013).
Correlations of ethanol yields with Glc and Xyl content, or
Ara/Xyl ratios, highlight the importance of carbohydrate structural features for biomass recalcitrance, supported by glycome
profiling results. Strong XG–cellulose associations (Hayashi,
1989) concur with XG epitopes only being extensively removed
from the walls when using 4 M KOH. As XG is more abundant
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in stems, this may contribute to their higher biomass recalcitrance. Moreover, distinct immunolabelling patterns may influence cell wall deconstruction; for example, as cellulose and lignin
are lower in leaves, more abundant demethyl-esterified HG may
reinforce organ cohesiveness. Consequently, as pectins are among
C. phytofermentans primary polysaccharide targets (Lee et al.,
2012b), higher pectin content could make foliar biomass more
prone to degradation. Additionally, as HG demethylesterification promotes recalcitrance (Lionetti et al., 2010) and
methyl-esterified HG is progressively reduced in muro (Zhang &
Staehelin, 1992; Mohnen, 1999), more abundant demethylesterified HG in later harvests may contribute to lower ethanol
yields.
Conclusions
Key differences in miscanthus cell wall fine structures and composition were highlighted between organs and harvests. Between
harvests, stem cell wall compositional variability is primarily seen
in cellulose and lignin, but in leaves it is in easily extractable
matrix glycans. Harvest- and organ-specific abundance, distribution, composition and ornamentation of cell wall polymers affect
amenability to deconstruction differently. Despite higher stem
sugar contents, extracted carbohydrate and ethanol yields were
higher from leaves. As a result, the same lignocellulosic feedstock
may present widely distinct biorefining potential depending on
plant developmental stage and relative organ contributions to
total harvested biomass. Understanding how cell wall components interact with each other and altogether respond to biorefining approaches is vital to optimize lignocellulosic biomass
deconstruction. Thus, a holistic view of the cell wall is promoted,
considering that certain components have variable impacts on
recalcitrance depending on overall cell wall assembly. A detailed
exploration of factors underpinning recalcitrance will be the
future focus of this ongoing study.
Acknowledgements
This work was supported by European Regional Development
Funding through the Welsh Government for BEACON, grant
number 8056; the Biotechnology and Biological Sciences
Research Council (BBSRC) Institute Strategic Programme Grant
on Energy Grasses & Biorefining (BBS/E/W/10963A01); and
the Office of Science (BER) Department of Energy, grant
DE-SC0006621. Glycome profiling studies were supported by
the BioEnergy Science Center (BESC), Oak Ridge National Laboratory, and funded by a grant (DE-AC05-00OR22725) from
the Office of Biological and Environmental Research, Office of
Science, U.S. Department of Energy. Generation of the CCRC
series of monoclonal antibodies used in this work was supported
by the NSF Plant Genome Program (DBI-0421683 and IOS0923992). We acknowledge BBSRC ‘Sparking Impact’ funding
and a Society for Experimental Biology Travel Fund to enable a
research visit of R.M.F.d.C. to the CCRC, University of
Georgia.
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Author contributions
R.M.F.d.C., A.W. and M.B. planned and designed the research.
R.M.F.d.C. performed most experiments and all data analyses.
Glycome profiling and immunohistochemistry were performed
in collaboration with Dr Ronald Clay, U.A. and S.P., under the
supervision of M.G.H. S.J.L. performed the Clostridium bioassays under the supervision of S.P.H. The manuscript was produced by R.M.F.d.C. and revised by S.P., U.A., S.J.L., S.P.H.,
A.W., M.G.H. and M.B.
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Supporting Information
Additional Supporting Information may be found online in the
Supporting Information tab for this article:
Fig. S1 Distribution of measurements of the minor monosaccharides fucose (Fuc) and galactose (Gal) released upon acid hydrolysis of miscanthus CWM.
Fig. S2 Distribution of measurements of the major monosaccharides arabinose (Ara), glucose (Glc) and xylose (Xyl) released
upon acid hydrolysis of miscanthus CWM.
Fig. S3 Distribution of measurements of the arabinose to xylose
ratio (Ara/Xyl) of miscanthus CWM.
Fig. S4 Distribution of measurements of released acetate upon
0.1 M KOH treatment of miscanthus CWM.
Fig. S5 Distribution of measurements of ferulic (FA) and
p-coumaric (pCA) acid released upon 1 M KOH treatment of
miscanthus CWM.
Fig. S6 Total carbohydrate recovered from each sequential
extraction step g–1 purified cell wall material (mg g 1 CWM)
estimated by the phenol-sulphuric acid assay.
Fig. S7 Heat map of the standard deviations (SD) from the mean
binding intensities shown in Fig. 2.
Fig. S8 Immunofluorescent labelling of cell wall glycan epitopes
in transverse sections from leaves and stems from M. 9 giganteus
(gig01) with CCRC-M174 (galactomannan-2) and before a base
treatment with 0.1 M KOH for CCRC-M155 (xylan-5, Me-Glc
substituted xylan).
Fig. S9 Mean binding values to different classes of cell wall glycan epitopes released at sequential extraction steps from leaf and
stem samples from miscanthus biomass at three developmental
stages (same data as in Fig. 5, but organized by organ).
Fig. S10 Principal components analysis of glycome profiling data
(data are presented for all samples from the six fractions obtained
during the sequential extraction).
Fig. S11 Principal components analysis of glycome profiling data
(data is presented for each individual extraction step performed
during the sequential extraction).
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Fig. S12 Principal components analysis of glycome profiling
(data presented independently for each organ).
Table S1 Listing of plant cell wall glycan-directed monoclonal
antibodies (mAbs) used in the glycome profiling screening
Table S2 Amount of carbohydrate recovered at each extraction
step g–1 of isolated cell wall material (mg g 1 CWM) based on
phenol-sulphuric acid assay for total sugar estimation
Research 1725
Table S4 Pearson coefficients and associated probability (P) of
the correlations between the results obtained for the
C. phytofermentans-mediated digestibility assessment assay and
several cell wall features
Please note: Wiley Blackwell are not responsible for the content
or functionality of any Supporting Information supplied by the
authors. Any queries (other than missing material) should be
directed to the New Phytologist Central Office.
Table S3 Monosaccharide and acetyl bromide soluble lignin contents of the residue left after the sequential extraction
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